Hoy: Myths of Managing Resistance 385 INTRODUCTION TO THE … · 2010. 3. 4. · Leibee and...

Hoy: Myths of Managing Resistance

385

INTRODUCTION TO THE SYMPOSIUM,THE MYTHS OF MANAGING RESISTANCE

M

ARJORIE

A. H

OY

Entomology & Nematology Department, University of Florida,Gainesville, FL 32611-0620

Resistance to pesticides in arthropod pests is a serious and increasingly perplexingproblem in Florida, the USA, and the world. Resistance to pesticides already has cre-ated significant economic, ecological, and public health problems in agricultural,household and garden, and medical/veterinary pest management programs. Exten-sive research has been conducted on diverse aspects of pesticide resistance, and wehave learned much during the past 40 years. However, to some degree, much of thediscussion about ‘resistance management’ has been based on ‘myths’. As an organizerof this symposium, one of my goals was to stress that managing resistance is a formi-dable task that will remain a perpetual pest management dilemma, because resis-tance is a fundamental survival response to stress by arthropods.

Five papers were presented in this symposium at the 1994 annual meeting of theFlorida Entomological Society, but one manuscript regarding the response by indus-try to resistance could not be published in this series.

In the first paper, Gary Leibee and John Capinera assess the impact of resistanceto pesticides in Florida and cite examples of resistance that limit pest managementoptions.

Julie Scott describes what we currently know about the molecular genetics of ar-thropod resistance to pesticides. The number of genes identified, and the diversity oftheir effects on the physiology of arthropods, verify that resistance is a normal re-sponse to diverse environmental stresses. ‘Pesticide resistance’ is part of a generalstress response with a long evolutionary history.

Leah Bauer describes what we know about resistance to various toxins of

Bacillusthuringiensis

(B.t.)

strains.

B.t.

provides microbial control of an increasingly diversegroup of arthropod species and is an increasingly important tool for integrated pestmanagement programs. The deployment of transgenic crop plants containing

B.t.

toxin genes is likely to be an effective method for inducing resistance in agriculturalpests. Despite the diversity of

B.t.

toxin genes isolated and cloned, cross resistancesare common. Thus,

B.t.

toxin genes are limited resources.Finally, I discuss a variety of resistance management methods and point out that

we cannot really

avoid

resistance—we can only

delay

its onset. I argue that resistancemanagement needs a paradigm shift that can best be accomplished if we recognizethat pest management must be changed from a single-tactic strategy to a multi-tacticmode. Delaying resistance, whether to traditional pesticides or to transgenic plantswith toxin genes, will require that we develop truly integrated pest management pro-grams, incorporating all appropriate tactics, including host plant resistance, culturalcontrols, biological controls, genetic controls, and biorational controls. Pesticidesshould be reserved for situations in which they perform best—as tools to resolve anunexpected pest population outbreak. Effective, fully-integrated IPM programs willdelay resistance because the number and rates of pesticide applications can be re-duced.

386

Florida Entomologist

78(3) September, 1995

PESTICIDE RESISTANCE IN FLORIDA INSECTS LIMITS MANAGEMENT OPTIONS

G

ARY

L. L

EIBEE

1

AND

J

OHN

L. C

APINERA

2

1

Department of Entomology and Nematology,University of Florida,

Central Florida Research and Education Center,Sanford, FL 32771

2

Department of Entomology and Nematology,University of Florida,Gainesville FL 32611

A

BSTRACT

Pesticide resistance in Florida was characterized through a survey and literaturereview. The survey was conducted in 1994 among public-sector entomologists to de-termine the current and future status, extent, context, pattern, and instances of pes-ticide (insecticide and acaricide) resistance in Florida. Results attested to the impactof pesticide resistance on the management of numerous arthropods in Florida.Twenty-five examples of insecticide and acaricide resistance were cited by survey re-spondents in agricultural, ornamental and landscape, medical and veterinary, orhousehold and structural pests. It remains possible to manage most arthropods by us-ing chemical pesticides, but the current and anticipated lack of efficacious materialsthreatens current practices in some areas. Trends in extent, context, or patterns of re-sistance were noted as follows: high value crops, frequently treated arthropods,smaller arthropods, and pyrethroids were all considered factors associated with resis-tance. Insecticide resistance and its management were reviewed in depth for the leaf-miner

Liriomyza trifolii

and the diamondback moth,

Plutella xylostella

, two majorinsect pests in Florida for which management options have become severely limitedbecause of insecticide resistance. Both cultural practices (continuous cropping, isola-tion, transport of infested seedlings) and pesticide use patterns (frequent applicationof broad spectrum pesticides) contributed to

L. trifolii

and

P. xylostella

resistance de-velopment. The history of pesticide resistance in these two insects is probably typicalof pest resistance in Florida and may portend similar future problems unless depen-dency on pesticides for pest suppression is reduced through adoption of IPM philoso-phy and practices.

Key Words: Insecticide resistance,

Liriomyza trifolii

,

Plutella xylostella

.

R

ESUMEN

La resistencia a los pesticidas en la Florida fue caracterizada a través de una en-cuesta y una revisión de la literatura. La encuesta fue conducida en 1994 entre los en-tomólogos del sector público para determinar el estado presente y futuro, extensión,contexto, patrón e instancias de la resistencia a pesticidas (insecticidas y acaricidas)en la Florida. Veinte y cinco ejemplos de resistencia a insecticidas y acaricidas fueroncitados por los que respondieron la encuesta sobre plagas agrícolas, de ornamentalesy de jardines, de importancia médica y veterinaria, o domésticas y de otras estructu-ras. Parece posible manejar la mayoría de los artrópodos usando pesticidas químicos,pero la falta actual y anticipada de materiales amenaza las prácticas presentes en al-gunas áreas. La tendencia en la extensión, contexto, o patrones de resistencia fuecomo sigue: cultivos de alto valor, artrópodos frecuentemente tratados, pequeños ar-trópodos, y piretroides fueron todos considerados como factores asociados con la resis-tencia.

Leibee and Capinera: Symposium on Pesticide Resistance

387

La resistenca a los insecticidas y su manejo fueron revisados en profundidad parael minador de las hojas

Liriomyza trifolii

y para la polilla de la col,

Plutella xylostella

,los insectos plagas principales en la Florida para los cuales las opciones de manejo sehan tornado severamente limitadas debido a la resisitencia a los insecticidas. Tantolas prácticas culturales (cosecha continua, aislamiento, transporte de plántulas infes-tadas) como los patrones de uso de pesticidas (aplicación frecuente de insecticidas deamplio espectro) contribuyeron al desarrollo de la resistencia de

L. trifolii

y

P. xylos-tella.

La historia de la resistencia a los pesticidas en estos dos insectos es probable-mente típica para la resistencia de las plagas en la Florida, y podría significarproblemas futuros similares a menos que la dependencia de los pesticidas para la su-presión de las plagas sea reducida a través de la adopción de filosofía y prácticas de

MIP.

Insecticide resistance has had an impact on the management of insect pests inFlorida since the mid-1940s following the widespread adoption of synthetic insecti-cides, especially the organochlorines, organophosphates, and pyrethroids. Numerousanecdotal reports exist, wherein consistently effective insecticides have become inef-fective and remained so for several seasons. Such reports have been considered ampleevidence of resistance development (Hoskins & Gordon 1956). In fact, Genung (1957)provided strong evidence based on anecdotal reports and data from field efficacy trialsfor resistance development in the cabbage looper,

Trichoplusia ni

Hubner, importedcabbageworm,

Artogeia rapae

(L.), a

Liriomyza

sp., and leafhoppers,

Empoasca

sp., ata session of the Florida State Horticultural Society Meeting in 1957 entitled “Sympo-sium-Vegetable Insect Resistance to Insecticides in Florida” (Brogdon 1957). Resis-tance episodes in Florida have also been documented in a number of species withlaboratory studies in which concentration-mortality response has been used to com-pare resistant and susceptible strains. Much of this work has been conducted withinthe last 10 years and involves species such as cabbage looper (Shelton & Soderlund1983), diamondback moth,

Plutella xylostella

(L.), (Leibee & Savage 1992a, Shelton etal. 1993, Yu & Nguyen 1992), silverleaf whitefly,

Bemisia argentifolii

Bellows & Per-ring, (G. L. L. unpublished data), house fly,

Musca domestica

L., (Bailey et al. 1970,Bloomcamp et al. 1987), German cockroach,

Blatella germanica

(L.), (Milio et al.1987, Koehler 1991, Hostetler & Brenner 1994),

Liriomyza trifolii

Burgess (Keil &Parella 1990, G. L. L. unpublished data), fall armyworm,

Spodoptera frugiperda

(J. E.Smith), (Pitre 1988, Yu 1992), cat flea,

Ctenocephalides felis

(Bouche), (El-Gazzar etal. 1986), and citrus rust mite,

Phyllocoptruta oleivora

(Ashmead), (Omoto et al.1994).

In hopes of providing a better understanding of current pesticide resistance and itsconsequences in Florida, we report here the results of a recent survey of public-sectorentomologists conducted to assess the extent of pesticide resistance in Florida, and itscurrent and potential impacts. In addition, we provide an in-depth account of two im-portant insect pests of vegetables in Florida, the dipterous leafminer

L. trifolii

andthe diamondback moth, for which management options have become extremely lim-ited because of insecticide resistance.

R

ESISTANCE

S

URVEY

During the spring of 1994, 16 public-sector entomologists were sent survey formsto measure their opinion about the extent of pesticide (defined as insecticide and ac-aricide) resistance in Florida, and its current and potential impact. We polled Univer-

388



sity and USDA entomologists from various backgrounds, representing the fields ofagricultural, ornamental and landscape, medical and veterinary, or household andstructural pest management. Entomologists with considerable field experience, and aclose relationship with producers or pest control professionals, were favored. We re-ceived responses from 14 of those surveyed, and 12 respondents provided useful infor-mation. Additional information was sought from other knowledgeable individuals toround out the survey. The questions and responses were as follows:

The Current and Future Status of Pesticide Resistance

Respondents were asked to indicate if resistance was: not a problem, a minor prob-lem, a significant problem, or a critical problem. Only a single response was re-quested. The time frame for future problems was specified as 10 years in the future.

The respondents differed in their assessment of the severity of the resistance prob-lem depending on the crop or environment being considered. Resistance was viewedto be a critical problem in greenhouses (foliage plants, flowering plants, and somewoody ornamentals), floriculture (both greenhouse and field-grown flowers), and ani-mal production (penned and free-ranging). Ornamental plants have long been consid-ered to be extremely sensitive to damage, hence they are treated frequently and proneto insecticide resistance problems. Resistance in animal production is a more recentphenomenon, however, apparently resulting from widespread use of insecticide im-pregnated ear tags.

Resistance was considered to be a significant, but not critical, problem in vegetablecrops, some field crops, and households. This might be viewed as surprising, becausemany vegetable crops, some field crops, and households in Florida receive insecticidetreatments at frequencies similar to the aforementioned situations where pesticideresistance was judged to be critical. It is likely that the severity of the problem is dueas much to corporate marketing strategies as to pesticide use patterns. Specifically,the pesticide market is smaller for greenhouse, floriculture and animal uses, so pes-ticide companies support fewer registrations. Therefore, when pesticide failures oc-cur, there are few options, or in some cases none. This, of course, results in a criticalsituation.

For medical pests, which in Florida is principally mosquitoes, the significance ofthe resistance problem apparently is related to location. Resistance was reported to bea significant problem in coastal locations, but only a minor problem in other areas.Coastal regions not only are extremely favorable for mosquito breeding, but a highproportion of the state’s population (79%) dwells along the coast, so there is frequentneed for chemical suppression.

Landscape plants seem to be relatively free of resistance problems. Woody orna-mentals are not usually planted in large single-species stands, which may help themto avoid development of high pest populations. Such landscape plants often tolerateconsiderable defoliation or pest density without obvious symptoms, so chemical treat-ment is not a regular feature of landscape maintenance. Also, in recent years therehas been a concerted effort to introduce native, hardy, pest-resistant plants into thelandscape, reducing the need for insecticide treatment. Among landscape plants, per-haps only turfgrass is treated regularly, and the southern chinch bug,

Blissus insu-laris

Barber, exhibits some degree of resistance, particularly in southern Florida.Nursery production of landscape ornamentals is also an exception, and mites canpresent resistance problems in this environment.

Although the number of pests displaying resistance to pesticides has increasedmarkedly in the last two decades, respondents generally did not see the resistanceproblem worsening greatly in the next 10 years. The only exception was the area of


389

household pest management, where the situation is anticipated to become critical.This generally optimistic attitude likely reflects faith in the agrichemical industry,which has continued to introduce novel pesticide chemistry or biorational materialsthat allow producers to continue with traditional agriculture and pest control prac-tices despite increasing numbers of pests that have become somewhat resistant to oneor more pesticides. The scientific community has also responded quickly and effec-tively to the onset of resistance by identifying alternative pest control chemicals andby helping to integrate other types of pest suppression into traditional production sys-tems.

The Extent of Resistance

Respondents were asked to indicate whether resistance applied to: a few com-pounds, numerous compounds, a few pests, or numerous pests. Up to two responseswere possible. Respondents were also asked to designate how many pests or com-pounds were affected and to indicate either a specific number or range.

The extent of the resistance problem was reported to be variable, depending onwhether the focus was the number of pests or pesticides. Resistance was generally re-ported to be limited to few pests in each commodity or environment. The number of re-sistant species was generally reported to be 3-5 per respondent, with a range given as1-10 per respondent. Although the number of species was small, the number of chem-ical compounds to which the pests were reported resistant was considerably larger.Respondents generally indicated that pests exhibiting resistance were resistant to5-10 compounds. The range in the number of compounds was given as from 1 productto all those on the market.

The Context of Pesticide Resistance

Respondents were asked to indicate if particular pests, crops, or environments ex-isted in which resistance occurred more frequently.

Respondents most frequently indicated that high value, damage-sensitive cropswere prone to have pesticide resistance problems. They cited greenhouse, floricul-tural, and vegetable crops as examples.

The environments next most frequently cited as having resistance problems werethose in which frequent or routine pesticide applications were made. Of course thiscorresponds to the aforementioned high value crops, but there are also situations inwhich value and damage sensitivity are not a major issue; examples are households,livestock, and certain field crops.

Only infrequently were the biological characteristics of the pests cited as favoringfrequent occurrence of resistance. Pests with short generation times and high intrin-sic rates of increase were suggested to be more prone to display resistance.

The Pattern of Pesticide Resistance

Respondents were asked if there were any patterns evident wherein entire classesof pesticide compounds or groups of arthropods displayed a tendency toward in-creased frequency of resistance, or whether resistance applied only to specific materi-als or pests.

Patterns of pesticide resistance related to chemical or biological taxon were not es-pecially evident to our respondents. Many said that pesticide resistance wasspecies-specific, that biological taxon was not a very good predictor of resistance prob-lems. A few, however, suggested that whiteflies, thrips, and especially mites were re-

390



sistance prone. Similarly, although cross resistance within chemical classes wasacknowledged, respondents indicated that they generally considered each pesticide tohave unique chemical properties, so that development of resistance was difficult topredict based on chemical taxon. The exception to this generalization seems to be thepyrethroids, where there is general acknowledgment that resistance is likely to de-velop.

Instances of Pesticide Resistance

Respondents were asked to name specific instances of pesticide resistance, includ-ing the pesticide, pest, and approximate date, and also to indicate whether the infor-mation on resistance was “documented” or anecdotal.

Instances of pesticide resistance in Florida provided by respondents are shown inTable 1. Surely this is not a complete list, either of pests or problem pesticides, butserves to demonstrate adequately the diversity of arthropod taxa affected. Also, ar-thropods found in numerous environments or crop systems are affected, and some his-torical trends are evident. Respondents acknowledged that only about one-half of thepurported cases of resistance are “documented,” with the remainder based on anec-dotal information. However, we carefully selected experienced entomologists andasked them to respond only in their area of expertise. Thus, we are confident that in-stances of misapplication and other potential sources of erroneous reports of resis-tance are not included. Because some of the “documented” resistance is from industrysources and not accessible to us, we have not included this specific information. Notealso that this table does not include information on the leafminer

L. trifolii

and the di-amondback moth, two insects with well-documented histories of insecticide resistancein Florida. A review of insecticide resistance in these two troublesome insects follows.

I

NSECTICIDE

R

ESISTANCE

IN

L.

TRIFOLII

Past and Present Situation

Prior to 1945, leafminer problems on celery and other vegetables in Florida wereapparently almost nonexistent. Control consisted mainly of clean-up measures andapplication of nicotine sulphate (Wolfenbarger 1947). Wolfenbarger (1947) recom-mended chlordane for control of leafminer on potatoes in south Florida. Harris (1962)reported that dimethoate, which was not labeled for celery, and diazinon and naledwhich were labeled, could control leafminer on celery in 1962. Genung et al. (1979) re-ported that with the use of diazinon, naled, and azinphos-methyl, the mortality of veg-etable seedlings and yield reductions declined and leafminer populations remainedlow until 1974, when they began to heavily infest celery and tomato. Genung et al.(1979) reported that in 1974 growers could not control leafminers on celery with diaz-inon, naled, or azinphos-methyl and that dimethoate, which was approved for use oncelery the same year, also did not give the desired level of control. Poe & Strandberg(1979) reported that oxamyl, which was approved for use on celery in 1975, was effec-tive for about two years. They also reported that in 1976 and 1977 leafminer on celerywas uncontrollable in Florida by any insecticide labeled for use. Florida growers ac-quired the use of methamidophos in 1977 and permethrin in 1978 for the control ofleafminer on celery. Permethrin became ineffective for leafminer control on celery inless than two years. Methamidophos was then considered the only insecticide thatgave any amount of control in celery in Florida, and it was considered marginally ef-fective. The possibility of effective chemical control did not come until the spring of


391

T

ABLE

1. E

XAMPLES

OF

I

NSECTICIDE

AND

A

CARICIDE

R

ESISTANCE

IN

F

LORIDA

C

ITED

BY

R

ESPONDENTS

IN

A

1994 S

URVEY

OF

P

UBLIC

S

ECTOR

E

NTOMOLOGISTS

.

ArthropodDate

(decade) Pesticide

House fly 1940 DDT

Musca domestica

(L.) 1950 chlordane, dieldrin, lindane, malathion1970 dimethoate, ronnel, tetrachlorvinphos1980 cyromazine, methomyl, various pyrethroids

German cockroach 1950 chlordane, dieldrin, lindane

Blatella germanica

(L.) 1960 allethrin, diazinon, malathion1970 carbaryl, propoxur1980 cyfluthrin, cypermethrin

Cat flea 1950-70 diazinon, malathion

Ctenocephalides felis

(Bouché)1970-801980-90

bendiocarb, carbaryl, propoxurcyfluthrin, fenvalerate, permethrin

Horn fly

Haematobia irritans

(L.)

1980 fenthion, fenvalerate, flucythrinate, permethrin, stirophos

Salt marsh mosquito 1950 DDT

Culex nigripalpus

1960 malathionTheobald 1990 methoprene

Soybean looper

Pseudoplusia includens

(Walker)

1970 acephate, methomyl, various pyrethroids

Fall armyworm

Spodoptera frugiperda

(J. E. Smith)

1970-80 malathion, carbaryl, methyl parathion, diazinon, trichlorfon, fluvalinate, bifenthrin, tralomethrin

Southern green stinkbug 1970 carbaryl, methomyl

Nezara viridula

(L.) 1980 endosulfanTobacco budworm

Heliothis virescens

(Fabricius)

1970 ethyl parathion

Corn earworm 1950-60 malathion, diazinon,

Helicoverpa zea

(L.) 1960-70 ethyl parathion, carbarylPepper weevil

Anthonomus eugenii

Cano

1990 fenvalerate, permethrin, oxamyl

Beet armyworm

Spodoptera exigua

(Hübner)

1980 chlorpyrifos, methomyl

Tomato pinworm 1970 carbaryl

Keiferia

1980 methomyl, fenvalerate

lycopersicella

(Walsingham)1990 oxamyl

Western flower thrips

Frankliniellaoccidentalis

(Pergande)

1980 pyrethroids

Mole crickets

Scapteriscus

spp.1970 chlordane

392



1982 when the celery industry secured the use of cyromazine. With the use of cyro-mazine, leafminer problems were considered under control until late 1989 when anunusual lack of efficacy occurred in the Everglades area.

Laboratory studies confirmed the presence of a high level of cyromazine resistancein a suspect strain of

L. trifolii

(G. L. L., unpublished data). Larval mortality in theresistant strain at 300 ppm of cyromazine, the highest label concentration used in thefield, was low enough to explain the loss of efficacy. The cyromazine resistance was ex-pressed as an incompletely recessive trait and not sex-linked. Backcrossing suggestedthat the resistance was conferred by a major gene. The resistance was considered un-stable since sensitivity returned in the resistant strain (from an LC

50

of about 440ppm to an LC

50

of about 85 ppm) within 5 generations of laboratory rearing withoutselection. This was consistent with a survey of leafminer populations that indicatedsusceptibility to cyromazine had returned during the summer of 1990 (J. S. Ferguson,unpublished data). This reversion was probably due to the immigration of susceptibleindividuals during what is traditionally a period of little or no celery production andvery little use of cyromazine.

The cyromazine-resistant strain was not resistant to abamectin (G. L. L., unpub-lished data), the only logical alternative insecticide available for control of leafminerin celery. This information contributed to the granting of a crisis exemption (Section18, FIFRA) in early 1990 for the use of abamectin in celery to control leafminer. Fur-ther efforts of the Florida Fruit and Vegetable Association, celery growers, CIBA,Merck Research Laboratories, and the University of Florida resulted in the subse-quent granting by the EPA (Section 18, FIFRA) in October 1990 of a specific exemp-

Green peach aphid

Myzus persicae

(Sulzer)1970-80 malathion, diazinon, oxydemeton-methyl,

dimethoateCabbage looper 1950-60 DDT, toxaphene, parathion

Trichoplusia ni

1960-70 endrin, mevinphos, naled(Hübner) 1970-80 methomyl

Cowpea curculio

Chalcodermus aeneus

Boheman

1980 endosulfan

Citrus rust mite

Phyllocoptruta oleivora

(Ashmead)

1990 dicofol

Yellow pecan aphids

Monellia caryella

(Fitch),

Monelliopispecanis

Bissell

1980 various pyrethroids

Silverleaf whitefly

Bemisia argentifolii

Bellows & Perring

1990 bifenthrin, fenvalerate, permethrin, endosulfan

Two-spotted spider mite 1980 fenbutatin-oxide

Tetranychus urticae

Koch1990 avermectin

Melon aphid

Aphis gossypii

Glover1990 acephate

TABLE 1. (CONT.) EXAMPLES OF INSECTICIDE AND ACARICIDE RESISTANCE IN FLORIDACITED BY RESPONDENTS IN A 1994 SURVEY OF PUBLIC SECTOR ENTOMOLOGISTS.

ArthropodDate

(decade) Pesticide

Leibee and Capinera: Symposium on Pesticide Resistance 393

tion for the use of abamectin in celery. Since then, abamectin has been used in celeryunder specific exemptions. These specific exemptions are unique in that, in order todiscourage the onset of resistance to abamectin, only two consecutive applications areallowed, forcing rotation with another insecticide. However, no other effective insec-ticide was available for rotation except for cyromazine which, due to reversion, hadbecome efficacious again. Since cyromazine and abamectin have different modes of ac-tion and no cross resistance was indicated, cyromazine was included in the leafminercontrol program under well-defined resistance management guidelines.

Management of Cyromazine Resistance in L. trifolii

A program for managing cyromazine resistance in L. trifolii was presented to cel-ery growers. The goal of this program was to control L. trifolii while increasing andpreserving susceptibility to cyromazine and minimizing the possibility of selecting forresistance to abamectin. Cyromazine use patterns and celery culture were suggestedthat would reduce selection of resistant phenotypes and encourage the immigration offeral, hopefully susceptible, leafminers into resistant populations.

Recommendations included: using noninfested transplants; initiating the sprayprogram based on a threshold to reduce the number of insecticide applications; start-ing with abamectin to maximize early control; rotating two sprays of abamectin withtwo sprays of cyromazine to avoid excessive use of one insecticide; finishing a plantingwith two applications of abamectin to reduce the number of adults emerging from thesoil and the trash after harvest; disking in trash as soon as possible to remove thissource of leafminers; and not using pyrethroids, such as permethrin and esfenvaler-ate, to minimize adverse effects on parasites and predators.

In addition, since acreage is very low in the production fields as harvesting ends(June) and the transplanting begins (September), it was recommended to not use cy-romazine during the summer (June through September) to prevent the continued se-lection of isolated populations and to encourage the immigration of susceptibleindividuals when celery acreage is at its lowest. Except for seedling production, Julyand August are otherwise free of celery. Not using cyromazine at all in seedling bedswas recommended, since transplanting from infested seedling beds is considered animportant mechanism for transferring resistant leafminers to the production fields.Lastly, seedlings were recommended to be grown distant from the field production ar-eas to reduce the chances of infestation by resistant leafminers.

INSECTICIDE RESISTANCE IN THE DIAMONDBACK MOTH

Past and Present Situation

Historically, the diamondback moth was considered a minor pest, usually includedin a complex of cabbage caterpillars along with the cabbage looper and the importedcabbageworm, Artogeia rapae (L.), but of much less importance. Control recommen-dations for the diamondback moth generally have been the same as for the other cab-bage caterpillars (Sanderson 1921, Metcalf & Flint 1939, Watson & Tissot 1942,Metcalf et al. 1951, 1962). Prior to the mid-1940s, insecticides used for cabbage cater-pillar control included nicotine, arsenicals, pyrethrum, rotenone, kerosene, and hotwater (150°F), and from the mid-1940s through the 1970s included DDT, toxaphene,parathion, methoxychlor, mevinphos, endosulfan, naled, methomyl, and methami-dophos. Bacillus thuringiensis was also available, but was not used extensively due toexpense and the perception of less than desirable control. In the early 1980s growers

394 Florida Entomologist 78(3) September, 1995

switched to the newly available and extremely effective pyrethroids, permethrin andfenvalerate for control of the cabbage looper and diamondback moth, both of whichhad become difficult to control with the other insecticides.

Insecticide resistance had long been suspected as the cause of the poor cabbagelooper control (Genung 1957, Workman & Greene 1970), and Shelton & Soderlund(1983) showed that a population from Florida was one of the most resistant to meth-omyl in the eastern U.S. The poor control of diamondback moth has been attributedto the destruction of parasites by excessive use of insecticides, such as methomyl,which were applied for cabbage looper suppression, but which were relatively ineffec-tive on diamondback moth. However, the poor control of diamondback moth may haveactually been the earliest indications of resistance problems.

Permethrin and fenvalerate proved to be very effective for control of all cabbage in-sects until the mid-1980s when growers observed that these insecticides were nolonger providing effective control of the diamondback moth. University trials reflectedthe same lack of control with fenvalerate (Leibee 1986) and from the winter of 1986-87 to the present, pyrethroid insecticides provided poor control of diamondback mothat Sanford, FL (G. L. L., unpublished data). Magaro & Edelson (1990) noted that fail-ures to control diamondback moth in south Texas were first reported by cabbage pro-ducers in the spring of 1987. Leibee & Savage (1992b) reported a high level ofresistance to fenvalerate in a laboratory strain of diamondback moth collected in cen-tral Florida in 1987.

Loss of efficacy with pyrethroids for control of diamondback moth caused growersto switch to intensive use of several organophosphates, endosulfan, and B. thuring-iensis subspecies kurstaki (Btk), all of which did not provide the level of control pro-vided by pyrethroids prior to resistance. At present, many diamondback mothpopulations have become very difficult to control with any of the registered syntheticinsecticides and Btk.

The presence of Btk resistance in Florida was immediately suspected because Btkresistance in diamondback moth had been reported in Hawaii (Tabashnik et al. 1990),Japan (Tanaka & Kimura 1991), and Malaysia (Syed 1992); it was eventually con-firmed for Florida (Leibee & Savage 1992a, Shelton et al. 1993). With the presence ofBtk-resistance, there were essentially no effective insecticides available for control ofmany diamondback moth populations in Florida until the recent introduction of B.thuringiensis subspecies aizawai (Bta)-based insecticides. Bta-based insecticides(those possessing the Cry1C toxin) are being successfully used in areas where Btk-based insecticides have failed. Lack of resistance to Bta in diamondback moth resis-tant to Btk has been documented in Japan (Hama et al. 1992), Malaysia (Syed 1992),and Florida (Leibee & Savage 1992a, Shelton et al. 1993).

Diamondback moth abundance has been considered low for several seasons in cen-tral Florida (G. L. L., personal observation). This is due in part to the return of sub-stantial amounts of natural control from parasites, which in turn is attributed toreduced pyrethroid use. Growers are not spraying as frequently for diamondbackmoth and are able to use Btk-based insecticides, suggesting a return of susceptibilityto Btk.

Management of Resistance in the Diamondback Moth

Crop culture and control recommendations were made that would reduce the se-lection of resistant phenotypes of diamondback moth and encourage the immigrationof feral, hopefully susceptible, individuals into resistant populations. These recom-mendations were based on the following knowledge. Susceptibility to Btk had beengreatly reduced in some populations. Bacillus thuringiensis resistance in diamond-


back moth in Hawaii was shown to be inherited as a recessive trait (Tabashnik et al.1992) and observations from field and laboratory studies in Florida suggested thesame (G. L. L., unpublished data). Bacillus thuringiensis subspecies aizawai-basedproducts (those possessing the Cry1C toxin) appeared to be effective in populationswhere Btk susceptibility was reduced. Tank-mixing Bt with mevinphos was shown tobe quite effective at reducing infestations in early season (G. L. L. unpublished data);however, the use of mevinphos was to be discontinued in 1995, eliminating the mosteffective insecticide other than Bt for diamondback moth control on cabbage in Flor-ida. Use of pyrethroids and carbamates can select for resistance that might further re-duce the efficacy of organophosphate insecticides and endosulfan, and also destroy theparasites and predators providing natural control of the diamondback moth.

Crop culture recommendations included: not growing cabbage in the warmestmonths (May through September in central Florida) when insect pressure is the high-est and Bt-based insecticides are the least efficacious; immediately disposing of cropresidues to prevent migration from heavily selected populations into new plantingsand seedling production areas; and using noninfested transplants, which not onlycontributes to control but also reduces spread of diamondback moths to new locations.Diamondback moths that infest purchased transplants may be highly resistant due toheavy usage of insecticides on the transplants or in fields near transplant productionareas. Producers growing their own transplants are at an advantage because theyhave more control over infestation levels and have specific knowledge about the resis-tance problems in their production areas.

Control recommendations included: inspecting crops frequently (about twice perwk) to determine the presence of the pest of concern or unexpected pests; beginninginspections at the seedling stage because reducing infestations in early season ap-pears to be critical to managing diamondback moth; minimizing insecticide applica-tions whenever possible by using action thresholds developed through research or byintuition; and using pheromone traps to monitor the presence or absence of diamond-back moth before and during the growing season, and also for monitoring peaks ofadult activity (Baker et al. 1982) for timing insecticide applications.

Specific insecticide recommendations included: using Btk and Bta as the principleinsecticides for control of diamondback moth; if the population was known to be sus-ceptible to Btk, alternating a Bta-based product with a Btk-based product to avoid re-petitive applications of the same insecticide to reduce the selection of resistance toany one product; using only Bta to insure maximum control if Btk resistance wasknown to be present or the status of Btk susceptibility was unknown; applying Bttwice weekly and tank-mixing with mevinphos weekly, starting with the tank-mix tomaximize the control that is critical early in crop; including endosulfan, chlorpyrifos,and methamidophos as alternatives or substitutes for mevinphos in the tank-mixeswith Bt, especially endosulfan, since it belongs to a different chemical class thanmevinphos; avoiding the use of carbamates; and, not using pyrethroids.

OBSERVATIONS ON INSECTICIDE RESISTANCE IN L. TRIFOLII AND DIAMONDBACK MOTH

Probably the greatest factor contributing to the development of insecticide resis-tance in L. trifolii and diamondback moth was long term and frequent use of single in-secticides or classes of insecticides. However, celery and cabbage cultural practices inFlorida probably contribute to the rapidity and degree with which insecticide resis-tance develops in these two insects. Both crops are grown in relatively small and iso-lated areas, or pockets, of agricultural activity. In addition, both crops are grown insome form year round. This isolation and the lack of a substantial crop-free period re-sult in the containment and “cycling” of resistant populations. This results in the


same population being exposed to insecticides continually. In addition, the constantuse of insecticides removes susceptible individuals that may be immigrating into thepopulation, thus preventing the opportunity for reversion.

These factors are believed to be especially evident with the development of highlevels of resistance in the diamondback moth in the 1980s. Several changes in the pro-duction of cabbage in the 1980s contributed greatly to the “cycling” of resistant popu-lations in the production areas and the movement of resistant populations betweenproduction areas within and outside Florida. Among these changes was the lengthen-ing of the crucifer production period by harvesting later in the spring and transplant-ing earlier in the summer. Prior to this situation, much of the summer(June-September) in central Florida was basically a crucifer production-free periodand diamondback moth populations were very low. Crucifer production was thrustinto the warmer, drier parts of the year when diamondback moth became more of aproblem, resulting in increased use of insecticides and subsequent exposure of addi-tional generations to more selection pressure. With development of the containergrown transplant industry and the ability to grow transplants in the summer, a situ-ation was eventually created in which crucifers were continually produced through-out the year in the same localities, either as transplants or field crops, or both.Insecticide resistant diamondback moth could move from the fields to transplants inthe summer, and be redistributed back to the fields in late summer and early fall.Therefore, heavy populations of resistant diamondback moth were being perpetuatedlocally throughout the year and continually exposed to insecticides. Opportunities forthe return of susceptibility by the immigration of individuals with susceptible pheno-types was essentially eliminated.

Transplants probably were a major factor in the development of resistance prob-lems in diamondback moth on a national level during the 1980s; container-growntransplants were popular and a healthy containerized transplant industry developedin the south. Much of the transplant production in the south in the winter and springsupplied more northern growers with transplants to establish stands in the springwhen environmental conditions were not conducive to direct seeding. Transplantswere also produced during summer months in the north to facilitate establishment ofstands in the south when soil temperatures are too high for germination.

Transplant growers likely were oblivious to the fact that they were shipping resis-tant diamondback moth. They were probably controlling the later larval stages thatcause obvious damage and shipping what appeared to be uninfested plants. Adultsflying into the open-sided greenhouses from the field could maintain a supply of eggsand early mining instars before shipment. In addition, larvae have been observed tobe deep inside the bud of the transplant, out of reach of the insecticide and the eye ofthe grower. It is possible that avoidance of the insecticide deposits drives diamond-back moth larvae emerging from the leafmine into the bud.

Another aspect of the transplant industry that could have contributed to the de-velopment of resistance was that only a few large growers in areas of high levels of re-sistance produced most of the transplants used in Florida and the rest of the U.S. Thisgreatly increased the probability of many growers receiving transplants from areaswith resistant diamondback moth populations.

CONCLUSION

The results of the survey attest to the impact of insecticide resistance on past andpresent pest management in Florida. The review of resistance problems with L. tri-folii and diamondback moth illustrates how insecticide resistance can complicate themanagement of pestiferous arthropods. Insecticide resistance seems to be pervasive


in Florida, suggesting that we are not adequately considering the consequences of theway we use pesticides. Resistance management strategies should be integrated withnonchemical control whenever possible. In the case of the aforementioned L. trifoliiand diamondback moth problems, crop management (isolation and continuous crop-ping) were key factors in development of resistance.

In most cases, pesticides are the most efficient, easiest, and cheapest methods ofcontrolling pestiferous insects and mites. The number of pesticides available is dwin-dling rapidly, however, due to cancellation of registrations and lack of re-registration.As a result, the conservation of arthropod susceptibility to the remaining pesticides,and to newly developed pesticides, is becoming extremely important. Unfortunately,we have not made adequate effort to conserve susceptibility of arthropods to pesti-cides. The minimum effort should include development of baseline data that would al-low the investigation of potential resistance episodes in a timely manner. Conservingpesticide susceptibility takes the development of knowledge and a commitment fromthose responsible for producing, using, and conducting research on pesticides. Thus,educational efforts should be enhanced.

Something everyone can do to help manage resistance to any pesticide is to refineits use. Application of the correct amount and type of insecticide in a timely and effi-cient manner would help forestall the onset of resistance, particularly if nonchemicaltechniques could be used to reduce the numbers and frequency of application. This isbest accomplished following research on, and implementation of, IPM strategies. In-creasingly, pesticide resistance management must be considered an important com-ponent of IPM.

ACKNOWLEDGMENTS

We gratefully acknowledge the assistance of the following individuals who gavegenerously of their time to make the survey component of this paper possible: JoeFunderburk, Jerry Hogsette, Freddie Johnson, Joe Knapp, Phil Koehler, Russ Mizell,Charlie Morris, Lance Osborne, Jim Price, Dakshina Seal, Dave Schuster, PhilStansly, and Simon Yu. We are also grateful to Brett Highland for his critical reviewof the manuscript. Florida Agricultural Experiment Stations Journal Series No. R-04551.

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81.

Scott: Symposium on Pesticide Resistance

399

THE MOLECULAR GENETICS OF RESISTANCE:RESISTANCE AS A RESPONSE TO STRESS

J

ULIE

A. S

COTT

Knipling-Bushland USDA Livestock Insect Research LaboratoryAgricultural Research Service

Kerrville, TX 78028-9184

A

BSTRACT

In this overview of the molecular genetics of resistance, pesticides are regarded asone of the many environmental stresses against which insects must defend them-selves to survive. Examined at the genetic level, pesticide resistance appears to be apreadapted response to stress and not due to novel mutations caused by pesticide ex-posure. The genetic mutations—gene amplification, altered gene regulation, struc-tural alteration of a gene—which result in resistance are described and explained anda possible distribution mechanism of resistance genes is considered. Resistance mech-anisms, their associated biological processes and the types of genetic mutations asso-ciated with each are detailed. Finally, the potential of molecular technology for thedevelopment of novel methods to detect and monitor for resistance is examined andcompared to more traditional technology.

400



Key Words: Pesticide resistance, molecular, mutation, genetic, adaptation, stress, in-sect.

R

ESUMEN

En esta revisión de la genética molecular de la resistencia, los pesticidas son con-siderados como uno de los muchos estréses ambientales de los cuales los insectos de-ben defenderse para sobrevivir. Examinada a nivel genético, la resistencia a lospesticidas parece ser una respuesta preadaptada al estrés y no debida a nuevas mu-taciones causadas por la exposición a los pesticidas. Son descritas y explicadas las mu-taciones genéticas-amplificación genética, regulación génica alterada, alteraciónestructural de un gen-que producen la resistencia, y es considerado un posible meca-nismo de distribución de los genes de resistencia. Son detallados los mecanismos deresistencia, los procesos biológicos y los tipos de mutaciones asociados con cada uno deellos. Finalmente, es examinado y comparado a la tecnología más tradicional el poten-cial de la tecnología molecular para el desarrollo de nuevos métodos de deteccción y

monitoreo de la resistencia.

When we discuss pesticide resistance, we generally refer to our ability to control apest, not the pest’s ability to defend itself. Yet resistance is really a form of self-de-fense. To an insect, exposure to a pesticide is just one of the myriad of dangers whichmust be avoided in order to survive. In this sense, pesticide exposure may be de-scribed as an environmental stress and resistance as the overt expression of an in-sect’s natural response to that stress. Insects have been confronted with lethal andnonlethal stresses for as long as they have existed. And, they have evolved effectivedefense mechanisms to deal with these stresses. Their potential to adapt and developresistance to stress becomes most apparent when examined at the molecular geneticlevel. It is the objective of this paper to examine how and why insects adapt to stress,particularly pesticide exposure at the molecular genetic level and to emphasize thatresistance is a part of the normal response of insects to stress. In addition, the advan-tages and limitations of traditional and molecular technologies for monitoring for re-sistance are briefly compared. This paper is a general overview to introduce readersto concepts of molecular genetics and pesticide resistance. It is not a technical reviewof the molecular biology of specific resistance mechanisms.

S

TRESS

, R

ESISTANCE

AND

T

OLERANCE

: D

EFINITIONS

AND

I

NTERACTIONS

Before discussing the molecular genetics of resistance, some definition and discus-sion of the relationships of stress, resistance, and tolerance is required. Stress hasbeen broadly defined as “any environmental change that acts to reduce the fitness ofan organism” (Koehn & Bayne 1989). Stresses may have physical, biotic, and/or toxiccomponents (Fig. 1) which, in turn, may be acute, chronic, and/or seasonal and affectinsects at the community, population, and/or individual level. The deleterious effectsof excesses in temperature or humidity are self-evident. Exposure to ultraviolet radi-ation can influence trophic-level interactions of entire communities to the advantageof one population at the expense of another (Bothwell et al. 1994). Predation, parasit-ism, disease, inter- and intraspecies competition all act to determine the success orfailure of a population to occupy a particular ecological niche. Toxic components of theenvironment are also stresses that can affect populations and can be divided intothree groups: pollutants, which may be natural or artificial, pesticides, and plant al-


401

lelochemicals. Each of these stresses can affect insects differently, but all must be ad-equately dealt with for continued survival.

Resistance has been defined as the development of the ability in a strain of an or-ganism to tolerate doses of a toxicant which would prove lethal to the majority of in-dividuals in a normal (i.e. susceptible) population of the same species (Anonymous1957). This definition is somewhat imprecise, because it infers that resistance can de-velop in an individual either before or after exposure to a toxin, two very differentevents. Resistance is the phenotypic expression throughout a population of a herita-ble trait that was already expressed in at least some of the individuals in the popula-tion

prior

to exposure to a toxicant. The development of a measurable shift in apopulation’s susceptibility to a toxin is due to the specific selection of these pre-adapted individuals in the population, often over several generations, by exposure toamounts of toxicant which are sublethal to the pre-adapted individuals but may ormay not be sublethal to others in the population. For many years this adaptive eventwas difficult to understand; however, we now know that the toxic components of somepesticides are similar to ones present in plants (e.g. pyrethrum) and that the detoxi-fication systems that deal with these plant allelochemicals are the same systems thatdetoxify pesticides. Therefore insects which can detoxify certain plant allelochemicalswell are pre-adapted to detoxify and develop resistance to pesticides which have thesame mode of action as the allelochemicals even before the insects are ever exposedto the pesticides.

Resistance and tolerance are often used interchangeably in the literature and todefine one another; however, tolerance is also used to describe shifts in susceptibilitythat occur within a single generation

after

exposure to stress which is not expressedby succeeding generations until

after

exposure to a similar stress. This phenomenonis different from the one previously described. For example 6-day-old bollworms,

He-

Figure 1. Examples of environmental stresses which can act on living organismsand force individuals, populations, and entire communities to continuously adapt tonew and ever changing conditions in order to survive.

402



liothis zea

(Boddie), only detoxify relatively small amounts of methyl parathion, butat 12 days, they can detoxify 30-fold more methyl parathion. If gossypol, a toxic alle-lochemical present in cotton, is added to the diet, then 12-day-old larvae can detoxifynot 30-fold but 75-fold more methyl parathion. The presence of gossypol in the diet in-duced the larvae to produce more detoxifying enzymes (Muehleisen et al. 1989). Yet,the 6-day-old progeny of these individuals do not retain either the 30- or 75-fold in-crease in tolerance of their parents but must develop it over time and after exposureto gossypol. The ability to metabolize methyl parathion is inherent; the increase intolerance to it is not. The difference between resistance and tolerance can becomeblurred when a population is subjected to a strong selection pressure, such as chronicpesticide exposure, and the mechanism that is induced by the pesticide exposure toyield tolerance is also the mechanism that is specifically selected to yield resistance.Resistance, as it will be discussed here, refers to a decrease in susceptibility which isheritable and does not need to be induced before it can be expressed; however, expo-sure to a stress, such as an insecticide, may result in an increase in the expression ofthe resistance gene(s) which may or may not be heritable.

Two other terms which are often used and may be confused with one another arecross-resistance and multi-resistance. Cross-resistance is resistance to two or moreclasses of pesticides which occurs because the pesticides have the same, or very sim-ilar, modes of action. Organophosphate and carbamate pesticides intoxicate by simi-lar modes of action, and resistance to one usually results in resistance to the other.Multi-resistance refers to resistance to two or more classes of pesticides because of thecoexistence of two or more different resistance mechanisms. For example, a resistantinsect may have both metabolic resistance to organophosphates and target-site resis-tance to pyrethroids.

T

HE

W

AYS

AND

M

EANS

OF

A

LTERING

G

ENETIC

M

ATERIAL

It should be apparent that pre-exposure to plant allelochemicals can only help ex-plain those cases of resistance where the pesticide is rendered inactive by the samedetoxification mechanism. It does not explain resistance mechanisms which do notappear to be selected for by plant allelochemicals or seem to arise spontaneously or in-crease in amplitude after exposure to a pesticide. How can exposure of the parentalgeneration to stress result in their progeny being more resistant to that stress? Theanswer is that exposure to a stress causes the genetic material (i.e. the DNA) to be al-tered.

There appear to be three general types of alterations, i.e. mutations, that can occurand result in resistance (Fig. 2). A gene may be

amplified

so that instead of only hav-ing one copy of the gene, there are now many copies present in the DNA. If an insecthas ten copies of a gene, then it can make ten times as much product as an insect withonly one copy of the gene. If the amplified gene encodes for a detoxifying enzyme, thenthat insect can detoxify 10-fold as much toxicant as the insect with only one gene.

The expression of a gene may also be altered to yield resistance. In this case, thereis only one copy of the gene present in the mutated insect but that gene’s regulationis altered so that it produces more (or less) product compared to a susceptible individ-ual. For example, in a susceptible insect the gene to gene-product ratio may be 1:1 butin a resistant insect that ratio may be changed. The gene may be

up-regulated

to pro-duce more product, that is, the gene to gene-product ratio is now 1:10 or

down-regu-lated

to make less product (the ratio is now 1:0.1). When a pesticide is applied in itstoxic form, up-regulation of a detoxifying enzyme will increase resistance. When apro-insecticide, i.e. the material must be metabolized first in order to become toxic, isapplied, down-regulation of the metabolizing enzyme will increase resistance.


403

The third type of mutation that can result in resistance is a structural change ina gene which yields a corresponding structural change in its product. A single

pointmutation

, i.e. one nucleotide in the gene’s coding region is substituted with a differentnucleotide so that a different amino acid is encoded for at a specific position and thischange causes the gene product to have a different three-dimensional structure, canresult in resistance in several ways. It may decrease the ability of the insecticide tophysically bind to its site of action, or increase or decrease the gene product’s abilityto metabolize the insecticide. A structural change does not alter the quantity of theproduct made but alters the quality of the product made.

It is important to recognize that these alterations to an insect’s DNA do not createnew genes. They only affect pre-existing genes. The idea that exposure to a pesticidecauses resistance genes to be “created” has been debated periodically; however, nosubstantiating data have ever been proffered. It is much more likely that resistancegenes already exist in the pesticide-naive population at low frequency prior to selec-tion by a pesticide.

How resistance genes are spread throughout a population is not completely cer-tain. It is clear that pesticide exposure plays a key role. However, other factors mayalso be important in the spread of resistance genes. One hypothesis which has re-ceived considerable attention is that

transposable

or

mobile elements

play a signifi-cant role in some cases of gene amplification. Transposable or mobile elements (TEs)are discrete sections of DNA that can move to new chromosomal locations and prolif-erate at a higher frequency relative to other genomic sequences (i.e. more and morecopies of the TE are inserted into the genome) after they have moved. TEs can alsomove genes that were previously not mobile and whose functions are not related tothe transposition. The genes that are moved with the TEs are also replicated at ahigher frequency (Berg 1989). Gene amplifications may be initially distributedthroughout a population by TEs or a gene may be transposed to a new location whereits expression is altered to yield resistance. For example, it has been indirectly dem-onstrated in the laboratory that the transposition of

alleles

, alternative forms, of thegene

Met

results in insecticide resistance in

Drosophila

(Wilson & Turner 1992). But

Figure 2. Graphic representation of the types of genetic mutations which occurand cause resistance. (a) A gene is amplified to increase its number of copies in the ge-nome and consequently increase the amount of gene product made (b) the regulatoryexpression of a gene is altered to increase the amount of gene product made (note thatgene expression may also be altered to decrease the amount of product made) (c) thegenetic code is rewritten to produce a structurally different product.

404



so far there is absolutely no evidence that such an event has occurred in a field popu-lation.

R

ESISTANCE

M

ECHANISMS

AND

A

SSOCIATED

T

YPES

OF

M

UTATIONS

Interestingly, only specific types of genetic mutations appear to be associated withspecific resistance mechanisms (Table 1). The two types of mechanisms that causehigh levels of resistance are generally referred to as metabolic resistance and target-site insensitivity, respectively. Each of these consists of several biological mecha-nisms. Metabolic resistance can be divided into three principle enzyme systems: cyto-chrome P450 monooxygenases (P450s), nonspecific esterases, and glutathione S-transferases (GSTs). Components of each of these enzyme systems may be mutated toalter the detoxification of a pesticide.

Cytochrome P450s catalyze a variety of detoxification reactions in insects, includ-ing the hydroxylation of DDT, the epoxidation of cyclodienes, the aromatic hydroxyla-tion of the carbamates carbaryl and propoxur, and oxidation of phosphorothioates(Feyereisen et al. 1990). Given the variety of reactions stimulated by these enzymes,it is likely that several different P450 enzymes are present in any one insect and thatseveral alleles of each gene may exist. Such is the case for the mosquito

Anopheles al-bimanus

in which seventeen P450 genes were discovered (Scott et al. 1994) and forthe termites

Mastotermes darwiniensis

and

Coptotermes acinaciformis

in which mul-tiple isoenzymes of cytochrome P450s were detected biochemically (Haritos et al.1994). There is no evidence to suggest that P450 genes are amplified or structurallyaltered to yield insecticide resistance. But there are numerous examples of their ex-pression being altered by various substances (Rose et al. 1991, Jeong et al. 1992, Wax-man & Azaroff 1992, Snyder et al. 1993). By definition, if the expression of a P450 is

T

ABLE

1. T

HE

G

ENETIC

M

UTATIONS

A

SSOCIATED

WITH

E

NZYMES

AND

R

ECEPTORS

T

HAT

R

ESULT

IN

D

IFFERENT

T

YPES

OF

R

ESISTANCE

.

Associated Genetic Mutations

Types ofResistance

Gene Amplification

Altered Expression

Structural Change

Metabolic

P450 oxidases ND

1

+ NDEsterases + ND +GSTs ND + ?

Target site insensitivity

Acetylcholinesterase ND ND +GABA receptor ND ND +Sodium channel ND ? ?JH receptor ND ? ?

Other

Reduced penetration — — —Behavioral change — — —

1

ND = not detected; + = confirmed or strongly indicated; ? = implied but not confirmed; — = no data available.


405

altered to yield resistance, then susceptible and resistant strains will have quantita-tively different amounts of that P450. Three P450s have been demonstrated to beover-expressed by resistant insect strains: P450Lpr (Wheelock & Scott 1992),CYP6A1 (Cariño et al. 1994), and CYP6A2 (Waters et al. 1992), respectively. P450Lprhas been directly implicated as the major enzyme causing pyrethroid resistance inone strain of house fly (Wheelock & Scott 1992, Hatano & Scott 1993). CYP6A1 ap-pears to be a major cyclodiene-metabolizing enzyme in the house fly (Andersen et al.1994). The primary function of CYP6A2 has not been reported. A major handicap todetermining which P450s are involved in resistance is our inability to distinguish theactivities of individual uncharacterized P450s from each other with a high degree ofaccuracy. The activity of a specific P450 towards an insecticide must be determinedbecause its over-expression does not necessarily prove that it is the enzyme responsi-ble for resistance. The over-expression of CYP6A1 by the house fly is a case in point.CYP6A1 is over-expressed in a house fly strain that is resistant to DDT, organophos-phates, and carbamates but does not have significant resistance to cyclodienes. To de-termine which P450s are actually responsible for resistance, more specific substratesare needed.

Esterases are a large group of enzymes which metabolize a wide variety of sub-strates. All esterases are able to hydrolyze ester bonds in the presence of water. Sincemany insecticides, especially organophosphates and carbamates, contain ester bonds,it is not surprising that the mechanism of resistance in many cases is caused by ele-vated levels of esterases (Fournier et al. 1987, Field et al. 1988, Carlini et al. 1991,Kettermen et al. 1992, Chen & Sun 1994). Esterase levels can be elevated by eithergene amplification or altered gene expression. So far, no data have indicated that theexpression of esterase genes is altered to yield resistance, but the molecular charac-terization of esterases is limited to a very few insect species and this type of mutationcannot be discounted.

Considerable data show that certain non-specific esterase genes are amplified toyield resistance. The esterases which cause resistance in

Myzus persicae

Sulz. and

Culex

mosquitoes have been particularly well-studied. In these insects the resistantesterase genes are highly amplified and up to 250 copies of the same gene may befound in a single individual (Mouchès et al. 1986, Poirié et al. 1992). The more the es-terase genes are amplified, the greater the level of resistance that they provide (Fieldet al. 1988, Poirié et al.1992). This increase in resistance appears to be because the es-terases interact with the insecticides more readily than the insecticides’ own target.When the esterases are present in approximately an equal molar ratio to the insecti-cides, they are able to effectively sequester the insecticides and then slowly hydrolyzethe insecticides (Devonshire & Moore 1982, Ketterman et al. 1992, Karunaratne et al.1993).

How recently these esterases have been amplified in

Culex

populations has been amatter of some debate. One group has suggested that two esterases associated withresistance, A2 and B2, were amplified in a single event within the past forty years, i.e.since the use of organophosphate insecticides became widespread, and that A2 and B2have been distributed across three continents by migration since that event (Ray-mond et al. 1991). If this hypothesis is correct then all A2 and B2 genes should beidentical. But data show that they are not. Kinetic studies of the insecticidal interac-tion of purified A2 and B2 esterases found that different forms of each enzyme werepresent in a number of resistant strains (Ketterman at al. 1993), and three amino aciddifferences have been found between the two B2 genes that have been sequenced(Vaughan et al. 1995). Therefore, not all of the A2 and B2 genes are identical and ei-ther these genes were amplified in at least two separate events or they were amplified

406



once long ago and have since diverged. Examination of other amplified esterase genesfrom different

Culex

strains clearly reveals numerous differences at the molecularlevel and strongly suggests that multiple amplification events have occurred withthem (Vaughan et al. 1995).

In addition to amplification, esterases may be mutated to produce structurally dif-ferent enzymes which are able to metabolize insecticides more efficiently. In the Aus-tralian sheep blow fly

Lucilia cuprina

and the mosquito

C. tarsalis,

acarboxylesterase appears to be structurally altered in resistant populations to pro-duce high levels of resistance to malathion (Whyard et al. 1994a, 1994b). In neitherspecies is more of the enzyme produced. The difference between susceptible and resis-tant populations is strictly a qualitative difference in the enzyme produced. Whetheror not the carboxylesterases from the blow fly and mosquito are homologous to eachother will only be known through further molecular and biochemical analysis.

The final group of enzymes which may provide metabolic resistance are the GSTs.Both organophosphate and cyclodiene pesticides can be detoxified by GST pathways.These enzymes have been somewhat less studied at the biochemical and molecularlevel in insects than the P450s and esterases. Elevated GST levels are found in manyresistant insect strains (Motoyama & Dauterman 1975, Ottea & Plapp 1984, Aham-mad-Sahib et al. 1994, Hoffman & Fisher 1994) and increased GST activity is clearlythe underlying resistance mechanism in some cases (Kao & Sun 1991, Wang et al.1991, Prapanthadara et al. 1993). But in other resistant populations the increasedGST activity does not cause resistance (Bush et al. 1993, Hemingway et al. 1993, Ar-gentine et al. 1994). Both insecticides and plant allelochemicals induce increased GSTproduction (Yu 1992a, Lagadic et al. 1993, Leszczynski et al. 1994), and generalistplant feeders seem to rely on GST pathways more heavily than do specialist feedersto metabolize plant allelochemicals (Yu 1992b). Because GSTs can metabolize a widevariety of substances, increased GST activity may be part of a generalized compensa-tory change due to exposure to an environmental stress. How GST activity is in-creased has only been examined in a few Diptera. In dipterans, increased GSTactivity does not appear to be the result of gene amplification. In both the house flyand

Drosophila

, several GSTs contribute to resistance and their expression appears tohave been increased through a regulatory change (Wang et al. 1991, Cochrane et al.1992, Fournier et al. 1992). In addition, at least one resistance gene is structurally al-tered in

Drosophila

(Cochrane et al. 1992). Whether or not this structural change af-fects the resistance level in the flies remains to be determined. As numerous GSTshave been found in several insects, each of which appears to be encoded by a differentgene (Cochrane et al. 1992, Fournier et al. 1992, Baker et al. 1994), it is likely that re-sistance caused by GSTs is due to altered gene expression and/or structural changesand is not due to gene amplification in most, if not all, cases.

The second major resistance mechanism is target-site insensitivity, which refers toan alteration of the molecule(s) that directly interacts with the pesticide to reduce tox-icity. Both acetylcholinesterase (AChE) and the gamma-aminobutyric acid (GABA) re-ceptor are known targets of insecticides, and resistant alleles of each have been found.Voltage-gated sodium channels and the juvenile hormone (JH) receptor are putativetargets of insecticides. Their direct interaction with insecticides has not been con-firmed, but it is clearly evident that they play a key role in the intoxication process.

Acetylcholinesterase is the target site of both organophosphates and carbamates.These pesticides bind to AChE and prevent the enzyme from stopping the action of theneurotransmitter acetylcholine. Multiple forms of AChE that confer varying degreesof resistance have been found in a variety of arthropods (Nolan et al. 1972, Devonshire& Moore 1984, Pralavorio & Fournier 1992). In each case examined so far, the affinity


407

of AChE for the pesticide has been reduced. Neither gene amplification nor altered ex-pression of the gene encoding AChE has been detected. Instead, point mutations haveoccurred to structurally change the enzyme. Recently, Mutero et al. (1994) identifiedfive point mutations in

D. melanogaster

which are associated with reduced sensitivity.Several strains of resistant flies were found to have a combination of mutations. Indi-vidually, the mutations gave only low levels of resistance, but when several of themwere combined, high levels of resistance resulted. Mutero et al. (1994) hypothesizethat decreased sensitivity by AChE is the result of a combination of several muta-tions, each of which provides a little resistance instead of the appearance of a singlemutation which yields strong resistance.

GABA receptors are the primary target of cyclodiene insecticides. In vertebrates,these receptors group together to form a complete chloride ion channel. It is inferredthat invertebrates have similar ion channels. Most cases of cyclodiene resistance ap-pear to be due to decreased sensitivity of the GABA subtype A receptor (ffrench-Con-stant et al. 1991), an integral part of the chloride ion channel. Like AChE, decreasedsensitivity by GABA receptors is due to a structural change of the protein. Neitheramplification nor altered expression of the GABA receptor gene has been detected.Unlike AChE, only a single point mutation which causes one specific amino acid to besubstituted with another results in high levels of resistance to cyclodienes. Otherpoint mutations have been detected, but no others appear to cause resistance or areconsistently associated with resistance (ffrench-Constant et al. 1993, Thompson et al1993).

Voltage-gated sodium channels play an integral role in the transmission of neuralimpulses. Pyrethroids disrupt neural transmissions by interrupting the normal func-tioning of voltage-gated sodium channels. Target-site insensitivity to pyrethroids, aphenotypic response commonly referred to as knockdown resistance (kdr), results inthese sodium channels becoming less sensitive to intoxication. Although it is clearthat sodium channels are adversely affected by pyrethroids, there is disagreement asto whether or not kdr is the result of a mutation to the sodium channels or to someother molecule which is integral to the functioning of the sodium channels. A pointmutation to a sodium channel, which could structurally alter it, was detected in a re-sistant insect strain (Amichot et al. 1992) but so far this mutation has not been shownto cause resistance. Other reports indicate that the mutation which causes kdr isclosely linked to (physically close to or a part of) the gene encoding one type of sodiumchannel (Knipple et al. 1994, Dong & Scott 1994) and this linkage has been inter-preted as strong evidence that a mutation(s) to the sodium channel gene is associatedwith kdr. On the other hand, a mutation to a regulatory protein or receptor could alsoresult in kdr. There is some indirect evidence to support this alternative hypothesis(Rossignol 1991, Osborne & Pepper 1992). A third hypothesis that has been proposedand for which there is limited electrophysiological evidence is that kdr is caused bychanges to two closely linked genes. One involves an altered sodium channel and theother may be associated with calcium-activated phosphorylation of a protein(s) in-volved with neurotransmitter release (Pepper & Osborne 1993). At this time, it canonly be stated that there is no indication that a gene amplification event is associatedwith kdr. It is most likely that either the expression of a gene associated with sodiumchannel function has been altered and/or a structural mutation to sodium channelsresults in kdr.

Juvenile hormone analogs such as methoprene compete with the natural hormonefor the JH receptor. Most resistance to JH analogs is either metabolic and/or a reduc-tion in penetration of the insecticide through the cuticle. The only reports of targetsite insensitivity to JH analogs are in mutants isolated from laboratory colonies of

408



Drosophila

(Shemshedini & Wilson 1990, Wilson & Turner 1992). The resistance gene

Met

that has been isolated from these colonies is associated with a less sensitive cys-tolic JH binding protein. Transposition of alleles of

Met

by a TE can induce resistance(Turner 1993); however, transposition is not required for the resistance gene to be ex-pressed. Therefore, it seems most likely that the insensitive JH binding protein is ei-ther the result of a mutation that structurally changes the protein or the expressionof the insect growth cycle has been altered. Gene amplification of JH receptors has notbeen implicated.

Two other types of resistance that have been described are the reduced penetra-tion of a pesticide and altered behavior to avoid a pesticide. It is presumed that the cu-ticular structure is somehow altered to reduce the rate of penetration of a pesticide.Avoidance behavior appears to be stimulated by brief contact, either through tactileor olfactory receptors, with a pesticide. Alone, neither of these mechanisms cause highlevels of resistance, but they are often found in combination with other types of resis-tance and can make a significant contribution to the overall resistance displayed byan insect. For example, Raymond et al. (1989) calculated that reduced penetration in-teracts with any other resistance mechanism multiplicatively. Experimental datasupport this conclusion (Hoyer & Plapp 1968, Plapp & Hoyer 1968). The underlyingphysiological, genetic and molecular mechanisms that cause these types of resistancecan only be speculated and the genetic mutations which cause them cannot be in-ferred at this time.

D

ETECTING

AND

M

ONITORING

FOR

R

ESISTANCE

Resistance is a widespread phenomenon and resistant populations of nearly alleconomically important pests can now be found (Georghiou 1994, Leibee & Capinerathis issue). Where control failures have occurred, the history of pesticide applicationusually indicates that chronic pesticide exposure resulted in high levels of resistancewhich caused the failures. Does the development of a resistant population then meanthat a control failure is inevitable? Certainly the continuous selection of the same re-sistance mechanism(s) over and over will result in resistance levels that are highenough to cause a control failure. But if the selection pressure (i.e. the pesticide) foreach resistance mechanism is removed prior to significant resistance developing, thena control failure may be avoided (for an alternative view on this subject, see Hoy thisissue). Keys to the successful avoidance of a control failure are the detection of resis-tance at low level(s), the routine monitoring for changes in the level(s) and type(s) ofresistance present in the pest population and the implementation of, and strict adher-ence to, a multi-tactic pest management program. In many cases, the two former ele-ments have not been fully utilized to allow for the design of an effective pestmanagement program. This lack of implementation is due in part to the unavailabil-ity of easy to use resistance detection methods.

The ideal detection method is fast, inexpensive, easy to use, diagnostic for all typesof resistance and able to detect resistance at frequencies as low as 1%. Numerousmethods to detect resistance are currently available but all fall short of being the idealtechnique. In fact, it is very unlikely that such an ideal technique will ever be devel-oped. Instead, we must rely upon traditional methods, principally bioassays and alimited number of biochemical assays, and novel molecular techniques to detect andmonitor for resistance. Certain general features are shared by the detection assayswithin each of these two technological groupings, i.e. traditional and molecular; andconsequently, they have similar advantages and disadvantages which are summa-rized in Table 2.


409

In general, detection methods which utilize molecular technology are better able todistinguish between the different resistance genotypes, i.e. heterozygotes (SR), ho-mozygous susceptible (SS) and homozygous resistant (RR), than traditional detectionassays. Because they detect only genetic differences, molecular assays can eliminatethe environmental components which often increase the variability in bioassay andbiochemical results. Direct comparison of a molecular assay and a bioassay (Aron-stein et al. 1994) indicates that molecular assays better approximate the perfectly di-agnostic assay described by Roush & Miller (1986) than traditional bioassays and,therefore, can require up to 5-fold fewer insects to yield the same information. And un-like traditional assays, molecular techniques can use material from a single insect toperform several different assays so that the resistance levels to a wide variety of pes-ticides can be determined from the same individuals. However, molecular assays arelimited to the detection of known resistance genes and a separate assay must be donefor each gene. Traditional bioassays are better able to detect the overall level of resis-tance present in a population in a single test. In fact, molecular assays cannot detectmany types of resistance at this time since with few exceptions we do not know whichspecific genes cause resistance. In time, molecular assays will be developed to detectmore resistance genes; however, it is unlikely that molecular assays will completelyreplace the traditional ones. In addition, molecular assays are more costly in both ma-terial and equipment and require greater technical training than simple bioassays.They cannot be used in the field and usually take significantly longer to complete.

Although m

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